Materials and methods for controlling vasculogenesis from endothelial colony forming cells

ABSTRACT

Materials and methods are disclosed for controlling vasculogenesis using building blocks of a collagen matrix and endothelial colony forming cells (ECFC). The building blocks may be isolated by fractionating an acid soluble Type I collagen. The building blocks comprising monomers and/or oligomers may be recombined in desired ratios to alter the matrix microenvironment and to influence ECFC behavior.

PRIORITY CLAIM

This application claims the benefit of U.S. Provisional PatentApplication No. 61/492,755 filed on Jun. 2, 2011 and incorporated hereinby reference in its entirety.

FIELD OF THE INVENTION

This invention relates generally to materials and methods forcontrolling vasculogenesis using for example some building blocks ofcollagen matrix, various cellular growth factors and endothelial colonyforming cells (ECFC).

BACKGROUND

Impaired vascular perfusion is a contributor to many disease states,including peripheral and cardiovascular disease, failure of tissue andorgan transplants, and impaired wound healing. The development of afunctional vascular network is necessary for development of clinicalscale tissue replacements to treat these disease states. Due to disease,defect and injury there is a need to promote the growth of newvasculature both in vivo and in vitro. When tissue is being developed,either in vivo or in vitro, for the treatment of a patient, there are avariety of circumstances under which it would be very useful to be ableto control the size, shape, bio-mechanical and biological properties ofthe tissue being developed. Development of a functional vascular networkis a major problem limiting current tissue engineering strategiestargeting repair and regeneration of damaged or diseased tissue.Recently endothelial colony forming cells (ECFCs) have been shown tovascularize a Type I collagen scaffold in vivo. ECFCs are the only cellsthat have been shown to possess direct in vivo vessel forming abilityupon transplantation. This has generated much interest in the use ofECFCs for tissue engineering strategies. However, there is still a greatneed for refinement of a defined microenvironment to locally deliverECFCs and guide vessel formation in vivo. Some aspects of the inventiondisclosed herein address this need.

SEQUENCE LISTING SEQ ID. NO. 1 CCACTACCAAGAAGGGATCTATCA; forwardprimer for ATP5B. SEQ ID. NO. 2 GGGCAGGGTCAGTCAAGTC; reverseprimer for ATP5B. SEQ ID. NO. 3 CATCGGAATATGTACCGACTGTT primerfor Cdc42. SEQ ID. NO. 4 TGCAGTATCAAAAAGTCCAAGAGTA reverseprimer for Cdc42. SEQ ID. NO. 5 CTGTCAGGAATGAGGATCTGAA primerfor MT1-MMP. SEQ ID. NO. 6 AGGGGTCACTGGAATGCTC reverseprimer for MT1-MMP. SEQ ID. NO. 7 CTGATCAGTTACACAACCAATGC primerfor Rac1. SEQ ID. NO. 8 CATTGGCAGAATAATTGTCAAAGA reverseprimer for Rac1.

BRIEF DESCRIPTION OF THE FIGURES

FIG. 1. Cartoon illustrating some steps in development blood vesselformation and some elements of the vascular system.

FIG. 2. Illustration of different forms of collagen isolated from nativecollagen by an acid extraction process.

FIG. 3. Graph of shear storage modulus (G′) versus collagenconcentration measured with collagen matrices comprised of varyinglevels of collagen monomer and oligiomer.

FIG. 4. Photomicrographs of collagen matrices comprised on differentlevels of AMW.

FIG. 5A. Graph of Fibril Density versus AMW (kDa).

FIG. 5B. Graph of G′ (Pa) versus AMW (kDa).

FIG. 5C. Graph of Ec (kPa) versus AMW (kDa).

FIG. 6. Photomicrographs of ECFCs seeded into collagen matrices.

FIG. 7A. Graph of Vacuole Area (mm²) versus Collagen Concentration(mg/ml).

FIG. 7B. Graph of Vacuole Area (mm²) versus Collagen Concentration(mg/ml).

FIG. 7C. Graph of Percent Vacuole (%) versus Vacuole Area (mm²).

FIG. 7D. Graph of Total Area (mm²) versus Collagen Concentration(mg/ml).

FIG. 8A. Graph of Vacuoles/mm² measured with monomers or oligomers.

FIG. 8B. Graph of Vacuole Area (μm²) measured with monomers oroligomers.

FIG. 8C. Graph of percent (%) Vacuoles>100 μm² measured with monomers oroligomers.

FIG. 8D. Graph of Total Area (μm²) measured with monomers or oligomers.

FIG. 9A. Graph of Vacuoles/mm² measured with monomers or oligomers.

FIG. 9B. Graph of Vacuole Area (μm²) measured with monomers or oligomers

FIG. 9C. Graph of Total Area (μm²) measured with monomers or oligomers.

FIG. 9D. Graph of Vacuoles/mm² measured with monomers,monomers/oligomers, or oligomers.

FIG. 9E. Graph Vacuole Area (μm²) measured with monomers,monomers/oligomers, or oligomers.

FIG. 9F. Graph of Total Area (μm²) measured with monomers,monomers/oligomers, or oligomers.

FIG. 10A. Graph of Vacuoles/mm² measured with monomers or oligomers at 3concentrations of collagen (mg/ml).

FIG. 10B. Graph of Vacuole Area (μm²) measured with monomers oroligomers at 3 concentrations of collagen (mg/ml).

FIG. 10C. Graph of Total Area (μm²) measured with monomers or oligomersat 3 concentrations of collagen (mg/ml).

FIG. 10D. Graph of Percent (%) Vacuoles measured with monomers oroligomers at versus Vacuole Area (μm²).

FIG. 10E. Graph Percent (%) Vacuoles measured with monomers, oroligomers versus Vacuole Area (μm²).

FIG. 10F. Graph Percent (%) Vacuoles measured with monomers, oroligomers versus Vacuole Area (μm²).

FIG. 11A. Photomicrograph of normal ECFC vacuole formation.

FIG. 11B. Photomicrograph of ECFC vacuole formation in the presence ofthe antibody MAB17781.

FIG. 11C. Photomicrograph of ECFC vacuole formation in the presence ofCasin.

FIG. 11D. Photomicrograph of ECFC vacuole formation in the presence ofthe antibody NSC23766.

FIG. 11E. Photomicrograph of ECFC vacuole formation in the presence ofTIMP-3.

FIG. 11F. Graph of Vacuoles/(mm²) measured with ECFC control and cellsexposed to MAB17781, Casin, NSC23766, or TIMP-3.

FIG. 11G. Graph of Vacuole Area (μm²) measured with ECFC control andcells exposed to MAB17781, Casin, NSC23766, or TIMP-3.

FIG. 11H. Graph of Total Area (μm²) measured with ECFC control and cellsexposed to MAB17781, Casin, NSC23766, or TIMP-3.

FIG. 12A. Photomicrograph of ECFC cultures grown on a tissue cultureplastic (Panel A) or on collagen matrices (Panel B).

FIG. 12B. Graph of Relative Expression to ATP-5B measured for thefollowing Cdc42, Rac1 and MT1-MMP.

FIG. 13A. Graph of Cdc-42 Expression/ATP-5B Expression measured foreither monomer or oligomer at both 0.5 and 30.0 mg/ml of Collagen.

FIG. 13B. Graph of Rac1 Expression/ATP-5B Expression measured eitherwith monomer or oligomer at both 0.5 and 30.0 mg/ml of Collagen.

FIG. 13C. Graph of MT1-MMP Expression/ATP-5B Expression measured witheither monomer or oligomer at both 0.5 and 30.0 mg/ml of Collagen.

FIG. 13D. Graph of Relative Expression/ATP-5B Expression versus Cdc-42,Rac1 and MT 1-MMP measured with either monomer or oligomer.

FIG. 14. Blot illustrating that FAK levels are elevated in ECFCs seededinto matricies polymerized from oligomer rich material.

FIG. 15. Photomicrographs (magnified 400×) showing that the RhoGTPases:RhoA, Rac1, and Cdc42 are active in vacuole formation in ECFCs.

FIG. 16A. Graph of Vacuoles/mm² measured with either 5e 5 or 2 e 6 ofECFC cells.

FIG. 16B. Graph of Vacuole Area (μm²) measured with either 5e 5 or 2 e 6of ECFC cells.

FIG. 16C. Graph of Total Area (μm²) measured with either 5e 5 or 2 e 6of ECFC cells.

FIG. 17A. Graph of Vacuoles/mm² measured with 5e5 or 2e6 ECFC cells andeither momomer or oligomer versus mg/ml of Collagen.

FIG. 17B. Graph of Vacuole Area (μm²) measured with 5e5 or 2e6 ECFCcells and either momomer or oligomer versus mg/ml of Collagen.

FIG. 17C. Graph of Total Area (μm²) measured with 5e5 or 2e6 ECFC cellsand either momomer or oligomer versus mg/ml of Collagen.

FIG. 18A. Graph of Percent (%) Vacuoles versus Vacuole Area (μm²)measured with either 5e5 or 2e6 ECFC cells in monomer rich matrices anda Collagen Level of 0.5 mg/ml.

FIG. 18B. Graph of Percent (%) Vacuoles versus Vacuole Area (μm²)measured with either 5e5 or 2e6 ECFC cells in monomer rich matrices anda Collagen Level of 3.0 mg/ml.

FIG. 18C. Graph of Percent (%) Vacuoles versus Vacuole Area (μm²)measured with either 5e5 or 2e6 ECFC cells in oligomer rich matrices anda Collagen Level of 0.5 mg/ml.

FIG. 18D. Graph of Percent (%) Vacuoles versus Vacuole Area (μm²)measured with either 5e5 or 2e6 ECFC cells in oligomer rich matrices anda Collagen Level of 3.0 mg/ml.

FIG. 19. Sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (4%)analysis (Lanes 1-3) and Western blot analysis with a collagen α1(I)antibody (Lanes 4-6) of a Type I collagen starting material, anoligomer-rich collagen fraction, and a monomer-rich collagen fraction.

DEFINTIONS

The following terms are defined as used herein, unless explicitly statedotherwise or clearly implied otherwise:

Collagen “monomers” represent single collagen molecules.

With respect to an acid soluble Type I collagen, a “monomer-richfraction” of the collagen is quantified as having an average molecularweight (AMW) of about 282 kDA.

Collagen “oligomers” represent covalently cross-linked monomers (e.g.,dimers=2 monomers, trimers=3 monomers, etc.).

Unless noted otherwise as used herein the acronym PSC referres to PigSkin Collagen.

With respect to an acid soluble Type I collagen, an “oligomer-richfraction” of the collagen is quantified as having an AMW of about 603kDA.

The term “about” refers to a range of values plus or minus 10 percent(e.g. about 1.0 encompasses values from 0.9 to 1.1).

SUMMARY

Some aspects of the invention include matrices that are suited forforming vessels comprising: a population of endothelial colony formingcells (ECFCs) in a polymerized matrix including at least one fraction ofan acid soluble Type I collagen. In some embodiments, these vessels maybe formed in vitro while in other embodiments they may be formed invivo, and in still other embodiments vessel development may begin invitro and continue after implantation in vivo.

In some embodiments, the fraction of the acid soluble Type I collagenthat is used to form the matrix is selected from the group consisting ofmonomers having an average molecular weight (AMW) of about 282 kDa andoligomers having an AMW of about 603 kDa. In some embodiments, at leastsome of the fraction of the acid soluble Type I collagen includes atelopeptide reactive aldehyde.

In some embodiments, the matrix further includes a second fraction ofthe acid soluble Type I collagen that differs from the at least onefraction. In some embodiments, the matrix is comprised of at least onefraction that is an oligomer-rich fraction and a second fraction that isa monomer-rich fraction. In some embodiments, the matrix includes moreof the oligomer-rich fraction than of the monomer-rich fraction. Theratios of the oligomer-rich fraction and the monomer-rich fraction maybe adjusted to create collagen matrices that have desirable physical andbiochemical properties and effects on the development of vacuoles inphysical contact with individual ECFCs or populations of ECFCs.

In some embodiments, one or more fractions of acid soluble Type Icollagen, or the total amount of acid soluble Type I collagen in amatrix, is present at a concentration of about 0.5 to about 3.0 mg/ml.In some embodiments, the population ECFCs is present in the matrix at aconcentration of about 5×10⁵ to about 2×10⁶ cells/ml.

Some aspects of the invention include methods of promoting vesselformation comprising the steps of: obtaining an engineered collagenmatrix from an oligomer-rich fraction of a Type I acid soluble collagen;and seeding the matrix with a plurality of ECFCs to form one or morevessels within the matrix.

In some embodiments, the matrix is richer in collagen oligomers thancollagen monomers. In some embodiments, the AMW of the collagen in thematrix is about 603 kDa. In some embodiments, the AMW of the collagen inthe matrix is greater than about 282 kDa. In some embodiments, the AMWof the collagen in the matrix is between about 282 kDa and about603+/−92 kDa.

In some embodiments, the matrix has a collagen content of about 0.5 toabout 3.0 mg/ml. In some embodiments, the collagen content of the matrixis about 0.5 mg/ml. In other embodiments, the collagen content in thematrix about 1.5 mg/ml. In still other embodiments, the collagen contentin the matrix is about 3.0 mg/ml.

In some embodiments, the methods of forming vessels include the step ofisolating acid soluble oligomers from Type I collagen.

In some embodiments, the matrix used to practice the invention isstiffer than a matrix formed using conventional Type I collagen.

In some embodiments, a stiffness of the matrix may be controlledindependently of a collagen concentration of the matrix.

In some embodiments, the matrix is in an unpolymerized state during theseeding step. In some embodiments, the seeding step comprises seedingthe matrix with about 5×10⁵ to about 2×10⁶ cells/ml.

Some embodiments include the step of implanting the matrix into adiseased area of a patient. In some embodiments, the matrix is implantedafter the seeded ECFCs begin to form vessels. In other embodiments, thematrix is implanted before the seeded ECFCs begin to form vessels. Thediseased area of the patient into which the matrix and the ECFCs or thematrix with vessels derived from the ECFCs includes an ischemic limb ofa human or animal patient in some embodiments.

By regulating cross link formation, a novel mechanism to alter thematrix microenvironment and influence endothelial cell behavior, Type Icollagen scaffolds can be engineered to support the formation of longlasting ECFC derived vessels. How collagen matrix microenvironment andphysical parameters (matrix stiffness, fibril density, and collagencross link composition) affect vascular network formation by ECFCs invitro was investigated. Further how collagen matrix design parametersmodulate a matrix-integrin-cytoskeleton signaling axis known to regulateendothelial cell lumen formation was investigated. The results provideinformation for the development of vascularized tissue constructs thatcan be controllably delivered to ischemic areas and improve the efficacyof human umbilical cord blood derived ECFC therapies for human subjects.

Collagen building blocks such as oligomers that include a reactivealdehyde and monomers that include a reactive aldehyde and theirreactive aldehyde free counterparts can be mixed in various combinationsto create a collagen matrix having select bio-mechanical properties.Properties affected by these combinations include the stiffness, matrixpore size, fibril density and the like.

Factors such as the degree of cross-linking and the concentration ofcollagen in a given mixture of collagen building blocks can be used toinfluence the fibril density and matrix stiffness of collagen-fibrilextracellular matrix. Such matrices can be seeded with ECFC and thejudicious use of various compounds that influence cell growth can beused to direct the growth of layers of these cells to form vascularstructure. Compounds that can be added to these matrices include, butare not limited to, MT1-MMP, α₂β₁-integrin, Rac 1, Cdc42, and the like.

As disclosed herein factors such as the density of the matrix may affectECFC vacuole formation. Similarly, the degree of cross linking of thematrix may also affect the degree of vacuole formation. Adjustingparameters that affect ECFC vacuole formation may influence the physicalproperties of lumen formed from by a given combination of collagenmatrix and ECFC.

Surprisingly, the degree of cross-linking in the collagen matrix altersthe ECFC mRNA express levels indicating its effect on cellular growthand development. For example, 2 dimensional versus 3-dimensional cellenvironments alter the cells' morphology and gene expression level andpattern. The activity of the enzyme Rho GTPases in vacuole formation isaffected by the matrix. This also appears to the same for the activityand expression levels of other cellular components such as Rac 1 andCdc42. These results are consistent with integrin cytoskeletonsignalling axis affect ECFC capillary morphogenesis.

The level of ECFC seeding within the matrix is positively correlatedwith the vacuole density of the cells and the bio-structures that theyform. Also, increasing the cell density requires that the rigidity ofthe matrix increase in order to form an increased vacuole areaassociated with the cells. This is also required in order to increasethe total area cell formed structures. Similarly, changing cell densityshifts the vacuole area distribution with the cell structures.

Vessels and vascular structures develop via vacuole formation withinindividual cells, coalescence of these vacuoles into multicellularlumens, and then remodeling via sprouting and branching to form aninterconnected network. Exemplary matrices of the present disclosuredemonstrated increased vacuole density, vacuole area, and total vacuolearea to support and enhance such vessel development.

Some aspects of the disclosure indicate that engineered vascularstructures are likely to require a rigid matrix with a low or moderatematrix fibril density (collagen concentration). Such matrices may resistcell traction and degradation while allowing for cell spreading andcell-cell interactions that promote capillary morphogenesis. Asdisclosed herein, the careful selection of collagen building blocks,ECFC levels and the application of compounds that effect cell growth canbe used to control the development of useful vascular structures.

DESCRIPTION

While the novel technology has been illustrated and described in detailin the figures and foregoing description, the same is to be consideredas illustrative and not restrictive in character, it being understoodthat only the preferred embodiments have been shown and described andthat all changes and modifications that come within the spirit of thenovel technology are desired to be protected. As well, while the noveltechnology was illustrated using specific examples, theoreticalarguments, accounts, and illustrations, these illustrations and theaccompanying discussion should by no means be interpreted as limitingthe technology. All patents, patent applications, and references totexts, scientific treatises, publications, and the like referenced inthis application are incorporated herein by reference in their entirety.

Early in development, a rapidly growing embryo exceeds a size that doesnot permit appropriate diffusion of nutrients or oxygen sufficientlydeep into the organism, requiring the development of a primitivevascular plexus. This process of de novo blood vessel formation known asvasculogenesis [3] allows for development of tissues beyond thediffusion limit of 100-200 μm [4, 5]. Referring now to FIG. 1, theprimitive vascular plexus and other vessels are continuously remodeledto accommodate growing or damaged tissues via sprouting andintussusceptive angiogenesis providing a system for transport of notonly oxygen and nutrients, but also cytokines and cells throughout theorganism. Briefly, during embryonic development mesodermal cells giverise to blood islands which consists of two cell types: angioblasts andhematopoietic elements. The blood islands are part of the primitivevascular plexus which is later remodeled by sprouting andintussusceptive angiogenesis. Recently EPCs have been shown to provideanother method of blood vessel formation in the adult via postnatalvasculogenesis.

This vascular system is lined by endothelial cells with a subjacentbasement membrane. Small caliber vessels are lined by a single layer ofperivascular cells, while larger more complex vessels have a wallcomprised of a complex extracellular matrix, nerves, and even smallervessels. The endothelium is crucial in maintaining normal vesselfunction. When the ability of the endothelium to repair or generate newvasculature is altered, the result is tissue damage and disease due toeither ischemia or inappropriate angiogenesis which can contribute totumor growth and metastasis. Hence an ability to understand themolecular mechanisms that govern vessel formation and remodeling is ofgreat interest for the treatment of these disease states [5, 6].

The remodeling of the vascular system to repair or generate new vesselsinvolves alterations to the surrounding matrix, cellular migration andproliferation, as well as tightly controlled signaling cascades. Asdisclosed herein, matrix scaffolds can modulate the in vitrovacuolization and in vivo vessel formation by human endothelialprogenitor cells (EPCs) by altering the biophysical environmentincluding mechanical and chemical properties.

Investigation into vasculogenesis and angiogenesis in matrix scaffoldsin vitro and in vivo [7-14] has often utilized mature endothelial cellpopulations such as human umbilical vein endothelial cells (HUVECs) andEPCs from umbilical cord and peripheral blood as well as from humanembryonic stem cells (hESCs). The formation of vascular networks fortherapeutic applications may require a population of endothelial cellswhich can be easily isolated, displays a high proliferative potential,and an ability to form vascular networks in vivo. While matureendothelial cell populations such as HUVECs have displayed the potentialto form functional vessels in vivo [11, 15, 16], they possess limitedproliferative capacity which will prohibit their use in large scaletissue constructs.

EPCs, are known to circulate in the bloodstream and home to sites inneed of vessel formation in both physiological and pathologicalsettings, have been examined over the past decade by numerousinvestigators for their therapeutic potential [17-20]. While studieshave demonstrated positive results in animal models, human trials haveresulted in mixed success [6, 18]. This may be due, in part, to severalfactors including: the rarity of the cells [21], controversy inisolation and expansion of EPCs [17, 19, 20, 22-24], and the use ofsystemic, rather than local delivery [6]. A major limitation has beenthe lack of a specific marker to identify an EPC and thus, greatheterogeneity in the types of cells that have been tested under theguise of an EPC.

During formation of the primitive vascular plexus, angioblasts, whichare precursors to endothelial cells [25] are thought to surroundemerging hematopoietic elements in close approximation [3]. Isolation ofputative EPC populations was originally based on cell surface antigensknown to be expressed on hematopoietic stem cells and endothelial cellsresulting in isolation of cells of both hematopoietic and endotheliallineages (reviewed in [26]). This method, first described by Asahara etal. [17], was later modified [19, 27] to deplete mature endothelialcells from culture to potentially enrich for EPC. Low densitymononuclear cells (MNCs) form adherent colonies, referred to ascolony-forming unit-Hill (CFU-Hill) after 5-9 days when plated onfibronectin coated tissue culture surfaces. CFU-Hill cells have beenshown to express cell surface antigens consistent with an endothelialcell phenotype, and ingest acetylated low-density lipoprotein (AcLDL), abehavior common to both endothelial cells and monocytes (Table 1).CFU-Hill cells also express several monocyte/macrophage cell surfaceantigens such as CD 14, CD45, and CD115, ingest bacteria, displaynonspecific esterase activity, and display limited proliferativepotential [8, 17, 28, 29]. Thus, the CFU-Hill assay does not identifyEPCs but rather permits enumeration of colonies of hematopoietic cells.While the hematopoietic cells do not directly contribute to theformation of new blood vessels (are not endothelium), these cells docontribute to neoangiogenesis via paracrine signaling pathways andfunction as circulating proangiogenic cells [30].

Recently another method of EPC isolation has identified a cellpopulation termed endothelial colony forming cells (ECFCs) [23] whichare also known as blood outgrowth endothelial cells (BOECs) [22, 31].Human umbilical cord or adult peripheral blood derived low density MNCsplated on Type I collagen coated tissue culture surfaces form adherentcolonies with a cobblestone morphology. These colonies first appear inculture between day 7 and 21, with cord blood derived colonies emergingearlier and at a higher frequency than adult blood derived colonies[23]. Ingram et al. developed a single cell assay to investigate theproliferative capacity of putative EPC populations. ECFCs demonstratedan ability to produce progeny in a clonal fashion, display a hierarchyof proliferative potential, and an ability to give rise to secondarycolonies when isolated from both umbilical cord and adult peripheralblood. Consistent with high proliferative behavior ECFC colonies exhibitrelatively high levels of telomerase [23]. While ECFCs express cellsurface antigens consistent with an endothelial cell phenotype [8, 23],they do not express hematopoietic or monocyte/macrophage cell surfacemarkers such as CD14, CD45, or CD115 [8] (Table 1).

TABLE 1 Phenotypic and functional characterization of CFU-Hill and ECFCs[8]. Von Willebrand factor (VWF), Ulex europaeus agglutinin 1 (UEA-1)lectin, acetylated low-density lipoprotein (AcLDL), vascular endothelialgrowth factor II receptor (KDR). Assay CFU-Hill ECFC Endothelial surfaceantigens CD31 92.31 ± 5.47 92.29 ± 1.32 CD105 74.36 ± 6.32 97.73 ± 1.79CD144 34.80 ± 8.74 99.15 ± 0.85 CD146  56.52 ± 10.00 94.21 ± 3.71 KDR99.19 ± 0.81  68.61 ± 11.26 WWF  67.21 ± 12.78 97.09 ± 2.05 UEA-1   41.± 11.67 100 AcLDL 73.68 ± 9.05 99.75 ± 0.25 Hematopoietic surfaceantigens CD45 98.15 ± 1.85  0.37 ± 0.37 CD44 98.53 ± 1.04  1.20 ± 0.74CD115 94.42 ± 2.52  0.28 ± 0.21 Macrophage properties Phagocytosis ofbacteria Yes No Nonspecific esterase activity Yes No Vasculogenicproperties Proliferative potential Some Robust Secondary colony-formingcapacity Some CFU-GM EC colonies In vivo vessel formation No Yes

One of the original defining concepts of an EPC was that of acirculating cell that possessed postnatal vasculogenic activity; theability to form a vascular system from a suspension of angioblast-likecells. ECFCs have displayed the potential to form blood vessels de novoin vivo when implanted in a Type I collagen matrix [8, 9] a Matrigelbased scaffold [10], or a hyaluronic acid based hydrogel [32]. No othercell type that has been referred to as an EPC can spontaneously form avasculature in vivo, though many of the hematopoietic derived cells arecapable of extravasating from the blood stream, migrating into a tissue,and attaching to any remnants of an endothelial basement membrane thatmay have persisted after endothelial dropout following cessation ofblood flow at a site of ischemia. Thus, the hematopoietic cells thatattach to the basement membrane remnant may appear to be forming avascular structure, but the cells are not synthesizing the matrix towhich they are attaching, a necessary step in stabilizing remodeledvasculature [33], and are therefore not endothelial cells. While, ECFCscould be isolated from peripheral blood to provide a patient specificcell source, adult blood derived ECFCs have a decreased proliferativepotential [23] and decreased ability to form functional vessels whenimplanted in a Type I collagen ECM [9] compared to umbilical cord bloodderived ECFCs. Thus, ECFCs display all of the properties of a cell thatone would envision as an EPC. Unfortunately, circulating ECFCs areextremely rare, being present at a frequency of 1/10⁶ cord blood and1/10⁸ adult peripheral blood mononuclear cells plated. Furthermore,there is no specific antigen that currently permits prospectiveisolation of the rare ECFC in the blood stream and discriminates thiscell from the occasional viable sloughed endothelial cells derived fromthe intima of systemic blood vessels.

Another opportunity exists to differentiate EPCs from hESCs. Endothelialcells derived from hESCs have shown the potential to form luminalstructures in vitro in both matrigel and Type I collagen scaffolds [34]and functional vessels when implanted in Type I collagen scaffolds invivo with murine 10T1/2 cells [35]. Additionally, vascular progenitorcells derived from hESCs were shown to form blood vessels when implantedwith and without hESC derived smooth muscle like cells in a Matrigelscaffold [36] Additionally, hESC seeded on to poly-(lactic-co-glycolicacid) scaffolds and transplanted between liver lobules ofimmunodeficient mice were extensively vacsularized by both host andhuman vessels, suggesting the in vivo differentiation of hESC into EPCs[37].

Additionally, the recent ability to reprogram adult differentiatedsomatic cells using a defined set of transcription factors to forminduced pluripotent cells (iPSCs) could provide a source of patientderived cells for vascularized tissue constructs [38-41]. Recent reportshave demonstrated endothelial cells can be differentiated fromfibroblast derived human iPSCs [42, 43] which could be a source ofautologous cells for angiogenic therapies. However, the proliferativepotential of both hESC and iPSC derived endothelial cells has not beenfully characterized. Further the ability of iPSC derived endothelialcell to form functional vessels in vivo has not yet been tested andfurther investigation is needed.

The native extracellular matrix (ECM) is a complex network of structuralproteins such as collagen, elastin, fibronectin, and proteoglycans [44].The engineering of such a complex matrix is very challenging and mostapproaches have used a simplified matrix as a model of the ECM [44]consisting of either synthetic or biological components. Syntheticmatrices used to study angiogenesis and vasculogenesis have been derivedfrom several polymers including polyglycolic acid (PGA), polylactic acid(PLA), polyethylene glycol (PEG) and others. Additionally, combinationsof polymers have been used to take advantage of specific properties ofeach component, such as poly (lactide-co-glycolide) (PLGA). Further,scaffolds from self assembling ionic peptides have been used to studyendothelial capillary network formation [45]. One advantage of thesesynthetic based scaffolds is the ability to fine tune microstructure anddegradation profiles by altering the processing and components of thescaffolds [14, 45].

Biological based matrices are typically composed of Type I collagen orfibrin as the main component [44]. Other materials used to generatebiological scaffolds to study angiogenesis and vasculogenesis includeMatrigel [10], elastin, hylauronic acid [32, 46], dextran, and alginate[47]. These biological based matrices have the advantage of beingbio-compatible, enzymatically degradable, and interacting with hostcells to promote vascularization. Scaffolds have additionally beenmodified to include growth factors [46, 47] and peptide sequences suchas arginine-glycine-aspartic acid (RGD) [47] to increase endothelialcell survival and adhesion.

While the ability to tune specific parameters in these scaffolds hasbeen more challenging than for synthetic based scaffolds, methods suchas altering source of matrix proteins, combining different biologicalproteins in scaffolds and cross linking matrix proteins providepotential tools to modulate the biophysical properties. Some studieshave demonstrated these effects on biophysical properties of collagenbased scaffolds. Collagen source and extraction method has been shown toaffect collagen fiber diameter and mechanical properties [48].Additionally, one report has demonstrated that the addition ofhyaluronic acid (HA) to the collagen based scaffolds alters thebiophysical properties [49]. Further, chemical fixations such asaldehydes, epoxides, and quinines as well as physical methods such as UVlight and dye-mediated photo oxidation can be used to cross linkcollagen fibrils in vitro [50]. Although these cross linking methods mayalter the bio-compatibility of the scaffolds. Finally, Baiely et al. [1]have described how altering the ratio of collagen oligomers and monomersmodulated matrix mechanical and microenvironment properties. Furtherinvestigation is needed to examine the role of cross linking of matrixcomponents and the addition of various proteins into biologicalscaffolds on the biophysical properties of these matrices.

Cells embedded in a 3D scaffold bind to and interact with the matrixcomponents. Cells seeded into a collagen based scaffold are able toreorganize the matrix [51, 52]. This interaction appears to occurbecause the collagen fibrils bind to integrin receptors which areanchored to the actin cytoskeleton [53]. Peptides such as RGD alsofacilitate scaffold adhesion to integrin receptors on embedded cells.Once bound the integrin receptors cluster and begin to form aggregatesof proteins such as talin, vinculin, and α-actinin known as focaladhesions [53, 54]. Focal adhesions serve as the entry point ofmechanical cues from the ECM into the cell and these cues influence cellshape, cell migration, cell survival, and cell differentiation [55-57].

The impact of the ECM on cell behavior has been demonstrated formesenchymal stem cells (MSCs). McBeath, et al., provided evidence thatmechanical parameters of the microenvironment dictate MSC shape andlineage commitment [58]. MSCs were plated on a PDMS(polydimethylsiloxane) micropatterned substrate which dictated theextent of cell spreading and shape. WMSCs that exhibited a spreadmorphology underwent osteogenic differentiation, while MSCs thatexhibited a rounded morphology underwent adipogenic differentiation.However, when the cytoskeletal tension was inhibited, MSCs underwentadipogenic differentiation independent of cell shape or morphology [58].

Ruiz and colleagues also demonstrated that force gradients can regulateMSC lineage commitment [59]. MSCs plated on a sinusoidal band wherecells cultured on the convex regions experienced high force, while cellsgrown on the concave regions were exposed to reduced forces. Cellscultured in regions of high force preferentially differentiated down anosteogenic lineage, while cells in the regions of low force underwentadiopogenic differentiation. Further osteogenic differentiation could beabrogated by inhibitors of cytoskeletal force generation [59].

Engler, et al., further demonstrated the impact of the microenvironmenton MSC behavior by showing that modulating matrix stiffness can directMSC lineage commitment [60]. Engler, et al., modulated matrix stiffnessby altering the concentration of bis-acrylamide cross linking Threematrix regimes were investigated: a low stiffness matrix thatapproximated the stiffness of brain, a medium stiffness matrix that wassimilar to the stiffness of muscle, and a high stiffness matrix that wasclose to the stiffness of osteoid. MSCs exhibited characteristics ofneurons, muscle cells, and osteocytes on the low, medium, and highstiffness matrix respectively. Further, morphology and lineage specificprotein expression of MSCs was dictated by the matrix stiffnesssupporting the importance of matrix stiffness on cell behavior [60].

The study of angiogenesis and vasculogenesis has evolved over severaldecades and includes both 2D and 3D assays. While the mechanism of tubeformation varies in different assays, all assays have demonstrated thatendothelial cell-matrix and cell-cell interactions are crucial for invitro endothelial cell network formation. One of the first in vitroassays was developed by Folkman and Haudenschild [61] in whichendothelial cells formed tubular networks on top of an endothelial cellmonolayer. The endothelial cell monolayer secreted its own matrixconsisting mainly of Type I collagen which was required for tubeformation [62].

In 2D in vitro endothelial cell vessel formation assays, the density ofmatrix proteins and endothelial cells influence the potential for tubeformation. Endothelial cells stimulated by fibroblast growth factor(FGF), known to induce endothelial spreading [63] and endothelial celltube formation in vitro [64], were plated on non adhesive plates withdifferent density coatings of fibronectin, gelatin, or Type IV collagen[65]. A high density of matrix proteins promoted cell spreading andgrowth, whereas a low density of matrix proteins resulted in cellrounding and death, but intermediate density resulted in tube formation.Tube formation could be induced at a high matrix protein density byplating endothelial cells at a higher cell density [65].

In 3D assays of in vitro capillary morphogenesis endothelial cells areseeded into a collagen solution so that the cells were evenlydistributed in the scaffold [66]. Many studies have investigated thecellular mechanisms of EC lumen formation in 3D Type I collagenscaffolds in vitro [11, 13, 66-76]. These studies have identified amatrix-integrin-cytoskeleton signaling axis that is critical in EC tubeformation [66, 69, 73, 75]. Matrix signals enter through integrinreceptors and activate Rho GTPase family members Rac 1 and Cdc42 toinitiate vacuole formation by embedded endothelial cells [75]. Vacuolesthen merge and coalesce to form multicellular structures. As thesecomplex structures are forming, the cells are also remodeling thesurrounding matrix. Membrane Type I-matrix metalloproteinase (MT1-MMP)dependent proteolysis allows the resident endothelial cells to createvascular guidance tunnels within the scaffold [77]. These channels, oncecreated, allow endothelial cell migration throughout the network ofguidance pathways. Interestingly, the integrin ligands which are engagedby the endothelial cells to interact with the scaffold switch duringthis process. Initially endothelial cells utilize α2β1 for vacuole andlumen formation in 3D collagen scaffolds [66]. As new matrix componentsare deposited, including fibronectin, laminins, nidogens, and Type IVcollagen, the endothelial cells upregulate expression of integrinstypically associated with these new matrix components and then use thereceptors for migration throughout the network of tunnels [77].

Endothelial cell behavior in 3D scaffolds is altered by the presence ofpericytes and other perivascular cells. In vivo vessel formation ofECFCs has been shown to be stabilized by mesenchymal progenitor cellsderived from either adult peripheral or umbilical cord blood leading toa longer time of persistence of ECFC derived vessels when implanted inMatrigel [78]. This effect has also been demonstrated in Type I collagenscaffolds. HUVEC derived in vivo vessels were stabilized by MSCsallowing the functional vessels to persist for greater than 130 days.Additionally these HUVEC-MSC composite vessels demonstrated avasoconstrictive response to endothelin-1 [79]. In vitro investigationshave suggested that this stabilization effect could be due to pericyteinteraction with endothelial cells within a scaffold inducingendothelial cell basement membrane deposition. Endothelial cells formvascular guidance channels which serve to recruit pericytes to the newlyformed vascular networks in vitro. Pericytes induce endothelial cells todeposit basement membrane proteins including fibronectin, laminin,nidogen-1, and perlecan. The deposition of basement membrane componentsserves to stabilize the vascular structures and diminishes remodeling invitro [33]. Understanding the role of accessory cells includingpericytes, CACs and others will be crucial to development of functionalvascular networks.

As outlined above, Endothelial Cell Network Formation studies havefocused on the importance of the matrix-integrin-cytoskeleton signalingaxis on capillary morphogenesis, only a few studies have investigatedthe effects of the mechanical properties of scaffolds on endothelialcell tube formation in vitro [70, 74, 80, 81]. Korff et al [70] seededType I collagen matrices with endothelial cell spheroids. When twospheroids were placed within 500-700 μm of each other, collagen fibrilswere induced to align along the axis between the spheroids. Further,endothelial spheroid sprouts would change direction towards otherspheroids, suggesting a role for the matrix of transmitting mechanicalsignals and aiding in the formation of multi cellular structures beforecell-cell contacts are formed. Additionally, soluble RGD peptides, whichinhibit collagen fibril-integrin binding, abrogated spheroid sprouting[70].

Sieminski et al. [74] seeded HUVECs or ECFCs into Type I collagenscaffolds with different collagen concentrations. The matrices wereeither left adhered to the well (adhered) or were released (freefloating) to further modify the mechanical properties of the scaffolds.Sieminski noted that changing the collagen concentration altered thematrix stiffness, ligand density, and had other biochemical effects[74].

The average lumen size and tube length of structures were dependent oncollagen concentration, endothelial cell type, and whether the matrixwas adherent or free floating. ECFCs seeded in 1.5 mg/ml collagenscaffolds that were adherent or free floating caused extreme contractionand cell death. ECFCs in 3 mg/ml collagen matrices formed tube likestructures. These structures were shorter and had wider lumens in 3mg/ml scaffolds that were adherent compared with those that were freefloating. HUVECs in 1.5 mg/ml matrices formed structures that weresimilar to those formed by ECFCs in free floating 3 mg/ml scaffolds,while structures formed by HUVECs in 3 mg/ml scaffolds had an appearancesimilar to ECFC structures in adherent 3 mg/ml scaffolds. The authorssuggest that the ratio of matrix stiffness and cell generated tensionare critical in the regulation of capillary morphogenesis [74].

Decreasing stiffness of an ionic self assembling peptide scaffoldincreases capillary network formation in vitro. HUVECs were seeded intoscaffold of stiffness ranging from 46-753 Pa. As stiffness decreased,elongation of capillary structures and the extent of single endothelialcells decreased. At the lowest stiffness tested multicellular capillarystructures were seen. Further decrease of scaffold stiffness below 46 Paled to compaction of the scaffold and did not permit endothelial cellnetwork formation. These results are similar to the results seen in TypeI collagen matrices, were compaction of gels at low concentration didnot permit for endothelial network formation [45]. These data provideevidence that the matrix microenvironment impacts EC tube formation invitro.

Scaffold contraction is dependent on cell type in Type I collagenmatrices. ECFCs were able to contract the matrices to a greater extentas compared to HUVECs suggesting ECFCs can generate a greater amount oftraction [74]. Sieminski speculated that the increased cell tractiongenerated by ECFCs could be due to increased expression of integrins,increased affinity for integrin ligands, increased sensitivity to FGF,or an increased sensitivity to phorbol esters [74].

Increased ECM density decreases capillary morphogenesis in fibrin basedscaffolds due to decreased diffusivity and cell generated traction [80,81]. HUVEC coated dextran beads were seeded into a fibrin scaffold witha monolayer of fibroblasts seeded on top of the matrix. Increasingmatrix density reduced sprouting and this negative effect could beovercome by seeding the fibroblasts within the matrix. The authorsdemonstrated an inverse relationship between matrix density anddiffusive transport of fluorescently conjugated mass markers. Theseresults suggest that decreased diffusion of cytokines could beresponsible in part for the decreased capillary morphogenesis withincreased matrix density [80]. Kniazeva et al. demonstrated thatchemical inhibitors of cell generated traction decreased sprouting ofHUVECs into the fibrin matrix and that this inhibition was directlyproportional the matrix density [81]. These results further support theidea that the ratio of cell traction and apparent matrix stiffness is animportant regulator of capillary morphogenesis.

Capillary morphogenesis occurs via vacuole formation within individualcells, coalescence of these vacuoles into multicellular lumens, and thenremodeling via sprouting and branching to form an interconnected network[90]. Several studies have identified cellular processes, the molecularpathway, and associated cytokines involved in this process [90,117-120]. From these studies it is evident that outside in signalingthrough a matrix-integrin-cytoskeleton signaling axis regulatescapillary morphogenesis in collagen based extracellular matrices. Theinteraction of the endothelial cells and the extracellular matrix duringvessel formation is initiated by integrin binding to collagen fibrils,resulting in integrin clustering, focal adhesion formation, cellgenerated traction and collagen fibril contraction and reorganization[89-93]. The remodeling of the ECM is resisted by the collagen fibrilnetwork. The resulting cell-matrix force balance determines the extentof capillary morphogenesis [1, 74, 121] suggesting that that thebiophysical and biochemical properties of the matrix are integral inregulating capillary morphogenesis.

While in vitro evidence suggests that physical properties of Type Icollagen matrices including fibril density and stiffness influenceendothelial cell capillary morphogenesis [70, 74, 99], the effect ofmatrix physical properties has only recently begun to be studied onendothelial colony forming cells (ECFCs) in vivo vessel formation.Collagen fibril density and stiffness impact human ECFC vasculogenesisin vivo [122] and Matrigel stiffness alters host capillary invasion[123]. Scaffolds with distinct biophysical environments were createdwith different shear storage moduli (stiffness) and fibril densities byaltering the collagen concentration. While human and total vasculardensity was greater in scaffolds with lower collagen concentrations andshear storage moduli, vascular area, which was dependent upon bothvessel size and density, increased with increasing collagenconcentration and shear storage modulus. [122]. Additionally, Matrigelwas implanted subcutaneously in mice with varied matrix stiffness(700-900 Pa), with 800 Pa scaffolds resulting in increased level of hostcapillary in growth [123]. While it is likely that complex processessuch as angiogenesis and vasculogenesis are regulated by a multitude offactors, these results suggest that increasing matrix stiffness andmatrix density results in increased vascularization in vivo.

While the above studies demonstrate a role of matrix stiffness andmatrix density in capillary morphogenesis, additional matrix propertiesare important in regulating tissue properties that have the potential toalter cell behavior and vessel formation. The ability of Type I collagento form the extracellular matrix of tissues with such distinct functionsas skin and bone is due largely to alterations in post translationalmodification and intermolecular-cross links. This can be seen by thedifferent collagen cross link chemistries that predominate in softtissue such as skin, sclera, and the cornea as opposed to stiff tissuessuch as bone, cartilage, and tendons [124]. Two enzymes lysylhydroxylase and lysyl oxidase regulate cross link formation. During theassembly of individual collagen monomers into fibrils, lysyl oxidasecatalyzes the formation of covalent cross links to form oligomers [125].

Referring now to FIG. 2, acid-extraction from tissues results in theisolation of both monomers (FIG. 2, Panel B) and cross linked oligomers(FIG. 2, Panel A). These molecules have reactive aldehydes resultingfrom acid labile divalent cross links, which will reform covalent crosslinks upon polymerization. Asterisks denote reactive aldehydes that formcovalent cross links and red lines denote native, non labile covalentcross links. These reactive aldehydes can be eliminated by chemicalreduction to alter the ability of these molecules to form cross linksupon polymerization (FIG. 2, Panels C-D). The ability to alter crosslink formation can be further inhibited by proteolytic digestion withpepsin to remove the telopeptide domains required for cross linkformation, resulting in atelo-oligomers and atelo-monomers (FIG. 2,Panel E).

The Voytik-Harbin laboratory has demonstrated that these unique collagenbuilding blocks can be used to polymerize matrices with varied G′ (Shearstorage modulus) versus collagen concentration profiles. Referring nowto FIG. 3, a graph is provided illustrating that collagen moleculesyield polymerized matrices with distinct relationships between Shearstorage modulus (G′) and collagen concentration. Oliogmer with reactivealdehyde (closed square), oligomer without reactive aldehyde (opensquare), monomer with reactive aldehyde (closed triangle), monomerwithout reactive aldehyde (open triangle), and pepsin digested oligomer(filled circle).

It has also been demonstrated that mixing monomer-rich and oligomer-richfractions in different proportions, which was quantified by averagepolymer molecular weight (AMW), alters the matrix microenvironment andphysical properties. Increase in AMW resulted in a decrease in matrixpore size (FIG. 4), but an absence of a positive correlation in fibrildiameter or fibril density (FIG. 5). Additionally AMW was positivelycorrelated with matrix stiffness (G′) and compressive modulus (Ec) (FIG.5) [1].

Referring now to FIG. 4, photomicrographs are provided illustrating theprojected pore size decreases with increasing AMW. All matrices werepolymerized with a collagen concentration of 0.7 mg/ml and under thesame conditions. 2D projections represent a total image thickness of 10μm (101 slices, scale bar=10 μm) [1].

In this study PSC oligomer-rich and monomer-rich fractions were used topolymerize matrices with altered biophysical properties to determine therole of collagen concentration, stiffness, and cross link chemistry onECFC vacuolization. Importantly this allows for an uncoupling of orindependent control over matrix stiffness and collagen concentration(fibril density), which has previously confounded investigations intotheir role in vascular network formation. Further the role of amatrix-integrin-cytoskeleton signaling axis previously demonstrated toregulate endothelial capillary morphogenesis in collagen based matricesin ECFC vacuole formation was tested and shown to be altered by thematrix microenvironment. Finally, the role of cell density in capillarymorphogenesis in matrices with varied microstructure and biophysicalproperties was examined and supports a role for the relevance of acell-matrix force balance in determining the extent of capillarymorphogenesis. The approach proposed here is the first to detail theseparate contributions of these two different matrix parameters inmodulating ECFC behavior in vitro. This will provide new informationthat will allow for a better understand of the underlying mechanisms ofECM-ECFC interactions. Importantly, this information is critical for thedesign and development of vascularized tissue constructs that can becontrollably delivered to ischemic or other diseased areas and improvethe efficacy of human umbilical cord blood derived ECFC therapies.

Therapeutic vasculogenesis, de novo blood vessel formation fromindividual endothelial cells, is a potential strategy to improve theseregenerative medicine therapies. Further ECFCs, an endothelialprogenitor cell population with clonal proliferative potential, havedemonstrated an ability to form functional vessels in vivo whenimplanted in collagen based scaffolds [8, 9, 78, 122]. However, thecellular response of ECFCs to biophysical and biochemical cues in theECM has only recently been explored.

Previous studies have demonstrated a role for matrix stiffness andmatrix density in capillary morphogenesis [81, 96, 122, 130]. Asreported herein there is a surprising role for both matrix stiffness andmatrix density the regulation of ECFC vacuole and lumen formation. Areport by Bailey, et al. [1] found a differential response of ECFCvacuole formation to matrices polymerized from monomer and oligomer richfractions. The instant disclosure demonstrates a role for collagen crosslink composition, by altering the oligomer content in collagen basedscaffolds, in modulating ECFC vacuole formation in vitro. By alteringthe oligomer content of the collagen scaffold the collagen concentrationand shear storage modulus (stiffness) could be independently varied,allowing their impact on ECFC vacuole formation to be investigated. Theprocess of capillary morphogenesis in this system was shown to dependupon a previously described matrix-integrin-cytoskeleton signaling axisinvolving integrin signaling and downstream RhoGTPase activity.Additionally alteration in collagen cross link composition was shown toalter both mRNA and protein expression by ECFCs during in vitro vacuoleformation and is the first to show alterations in FAK signaling duringvacuole formation. Finally cell density was shown to alter ECFC vacuoleformation dependent upon the matrix biophysical properties, supportingthe role for cell-matrix force balance in vessel formation.

ECFC vacuole density, vacuole area, and total vacuole area increasedwith increasing collagen concentration. This was in part due to a shiftin vacuole distribution toward larger vacuoles. These results areconsistent with previous reports that altered matrix density alters invitro capillary morphogenesis. However, previous reports have found thatincreased matrix density leads to decreased endothelial lumen formation.These differences could be due to the use of lower cell seeding density[74] and matrix composition such as peptide [45] or fibrin basedscaffolds [80, 81]. In the present study at a cell seeding density of5×10⁵ cells/ml ECFC total vacuole area decreased with increasingcollagen concentration. Additionally ECFCs have been shown to exhibitincreased matrix contraction and cell elongation compared to otherendothelial cell populations [130]. Thus the ECFCs may exert greatercell traction requiring more matrix resistance to drive vacuoleformation. It is possible that these different results based on cellseeding density, endothelial cell source, or matrix compositions betweenvarious studies are due to alterations in the cell-matrix force balance.

As disclosed in Bailey, et al. [1] that collagen cross link chemistryalters ECFC vacuole formation in vitro. As reported here in ECFCs seededinto matrices polymerized from oligomer rich fractions demonstratedincreased vacuole density, vacuole area, and total vacuole area comparedto ECFCs cultured in monomer rich matrices. This was due in part to analtered vacuole area distribution, with ECFCs cultured in oligomer richmatrices displaying a shift in the distribution toward larger vacuoleareas. Increased oligomer content in matrices polymerized with a matchedshear storage modulus (stiffness) of 200 Pa or matched collagenconcentration of 1.5 mg/ml resulted in increased ECFC vacuole densityand total vacuole area. Finally, oligomer rich and monomer rich matriceswith matched collagen concentrations of 0.5, 1.5, and 3.0 mg/ml werepolymerized and seeded with ECFC. At each collagen concentration theoligomer rich matrix demonstrated increased ECFC vacuole density andtotal vacuole area.

The differences in cell behavior in the different oligomer rich andmonomer rich matrices suggest the importance of multiple biophysicalproperties in regulating ECFC vacuole formation. Oligomer rich matriceshave a lower collagen concentration than monomer rich matrices with thesame stiffness. Additionally, oligomer rich matrices have a higherstiffness that monomer rich matrices polymerized with the same collagenconcentration. Together these data suggest that a rigid matrix with lowfibril density (collagen concentration) is supportive of ECFC vacuoleformation. The ability to generate these characteristics in a matrix isdependent upon the ability to modulate the collagen cross linkcomposition to increase interfibril branching which has been shown tocontribute to the matrix stiffness [131]. In this way the matrix is ableto resist cell traction and degradation, while allowing for cellspreading and cell-cell interactions necessary for capillarymorphogenesis to occur. These results further suggest that collagenderived from varied tissue sources which display different cross linkchemistries could be used to polymerize matrices with a wide range ofbiophysical cues to guide ECFC vacuole and lumen formation.

Previous work has detailed the role of a matrix-integrin-cytoskeletonsignaling axis in endothelial capillary morphogenesis. This axisinvolves integrin binding to collagen fibrils leading to downstreamRhoGTPase signaling and MMP mediated matrix degradation and remodeling.The use of specific chemical inhibitors of β01 integrin, Cdc42, Rac1,and MT1-MMP showed that this signaling axis is involved in ECFC vacuoleformation in vitro. While all inhibitors led to a reduction in totalvacuole area, the inhibition of Cdc42 led to reduction in total vacuolearea by decreasing vacuole density, while the inhibition of Rac1decreased total vacuole area by decreasing the average vacuole area.These results suggest that Cdc42 is intimately involved in theinitiation of vacuole formation, while Rac 1 is involved in theenlargement and remodeling of the vacuoles. Thus it is possible thatthese molecules could be used to either inhibit the initiation ofvasculogenesis or the continued remodeling of a primitive capillarynetwork. Further investigation into the nuanced role of these moleculesin vacuole formation particularly in vivo is required.

Endothelial cell behavior in traditional 2D tissue culture plastic isdifferent than in 3D environment seen in tissue scaffolds and in vivoenvironments. ECFC mRNA expression of Cdc42, Rac1, and MT1-MMP, threegenes known to play a role in capillary morphogenesis, were shown to beup regulated in 3D collagen based matrices compared to culture on 2Dtissue culture plastic. These differences are not unexpected based onthe differences in cell morphology in the different environments.Sacharidou et al. [128] have previously demonstrated that Cdc42 and MT1-MMP activity is necessary for endothelial cell migration throughcollagen matrices by not on 2D surfaces, suggesting that Cdc42 andMT1-MMP play a significant role in endothelial cell 3D but not 2Dbehavior. This may in part explain why the mRNA levels are up regulatedin 3D culture as ECFC begin to undergo capillary morphogenesis andrequire active Cdc42 and MT1-MMP to initiate the process.

In order to determine if collagen concentration and cross linkchemistry, which altered ECFC vacuole formation, altered cell signaling,RNA was isolated from ECFCs seeded in 3D matrices. While collagenconcentration did not demonstrate an effect on mRNA expression, collagencross link composition did alter the expression of Cdc42, Rac1, andMT1-MMP transcript levels. ECFCs cultured in oligomer rich matricesdisplayed increased expression of Cdc42 and MT1-MMP and reducedexpression of Rac 1 mRNA compared to ECFC cultured in monomer richscaffold. These data suggest that Cdc42 and MT1-MMP mRNA expression isincreased as the ECFC are undergoing capillary morphogenesis. Rac 1 mRNAexpression is increased in monomer rich matrices which yield reducedECFC vacuole formation. This increase in Rac1 expression may be theresult of the multiple roles of Rac1 in cell behavior [133]. Rac1 isknown to be involved in cell migration [133, 134] and the endothelialcells that do not undergo capillary morphogenesis do exhibit anelongated morphology. It is possible that in this environment the cellsmigrate throughout the matrix instead of initiating vacuole formationand that cell migration requires higher levels of Rac1 than does vacuoleformation. It is interesting to note that collagen concentration doesnot alter ECFC mRNA expression of Cdc42, Rac 1, and MT1-MMP even thoughit alters vacuole formations. One possible explanation for this is thatthere is a different level of active protein expressed in the differentcollagen concentrations. The FRET based RhoGTPases shown to be involvedin ECFC vacuole formation in this study represent an ideal system tofurther investigate the protein expression levels in capillarymorphogenesis at different collagen concentrations.

These results suggest that FAK protein levels are increased in ECFCcultured in oligomer rich matrices. FAK levels were shown to beincreased in oligomer rich matrices compared to monomer rich matrices ateither matched collagen concentration of 3.0 mg/ml or matched stiffnessof 200 Pa. Previous work has demonstrated a mechanotransductionsignaling pathway though β1 integrin and Src [135] to RhoGTPases duringcapillary morphogenesis. This data extends that signaling pathway toinclude FAK. It is not surprising that a matrix which is more supportiveof vacuole formation would increase FAK expression as this molecule isknown to be involved in focal adhesion formation and signaling withintegrin receptors to the cytoskeleton [136, 137]. It further suggestsFAK as a potential target to modulate vessel formation in futurestudies.

The results of this study have demonstrated a role for matrix stiffness,matrix density, and collagen cross link chemistry in regulation in vitrovacuole formation. Each of the variables plays an important role indetermining the microenvironment that the encapsulated endotheliumexperiences and responds to. Previous work has also demonstrated a rolefor cell-matrix force balance in capillary morphogenesis [74, 80]. Tobegin to test if the cell generated tension was also responsible foraltering vacuole formation, ECFCs were seeded in collagen scaffolds ateither 5×10⁵ cells/ml or 2×10⁶ cells/ml and vacuole formation wasanalyzed. While increasing cell density resulted in elevated vacuoledensity it reduced average vacuole area, and resulted in no differencein total vacuole area. While ECFCs seeded into matrices polymerized at0.5 and 3.0 mg/ml from monomer rich fractions and 0.5 mg/ml fromoligomer rich fractions demonstrated similar results, ECFCs seeded into3.0 mg/ml matrices demonstrated increased vacuole density, average area,and total area. Additionally the vacuole distribution for ECFCs seededinto the monomer and 0.5 mg/ml oligomer matrices displayed a shift tothe left, while ECFCs seeded into the 3.0 mg/ml oligomer matrixdisplayed no such shift. These data suggested the possibility that theratio of matrix stiffness and cell tension was being altered by theincreased cell density and that the monomer and 0.5 mg/ml oligomermatrices were unable to sufficiently resist cell traction and remodelingleading to stagnation in vacuole formation. This warrants additionalinvestigation possibly by directly altering the cytoskeletoncontractility using an agonist such as lysophosphatidic acid (LPA) or anantagonist such as Blebbistatin.

The results disclosed herein clearly show that the biophysicalproperties of collagen based matrices, specifically collagenconcentration, stiffness, and cross link composition modulate ECFCvacuole formation in vitro. Additional, results provided herein,demonstrate the role of cell matrix force balance in driving in vitrocapillary morphogenesis. Further demonstrating that the signalingcascade regulating this process is altered by changes in the matrixmicroenvironment. It is likely that all of the matrix design parametersdescribed here will impact in vivo vasculogenesis, and allow for thedevelopment of therapeutically relevant strategies to deliver ECFCs todamaged or diseased tissues.

Experimental Section Culturing of ECFC

Human umbilical cord blood derived ECFCs were isolated and cultured aspreviously described [23]. ECFCs were used between passage 5 and 10 forall experiments.

Separation of Collagen into Monomer-Rich and Oligomer-Rich Fractions

Type I collagen, comprising monomers and oligomers, was isolated frompig skin (PSC) and solubilized in acetic acid (0.005 M) to achieve adesired collagen concentration (4 mg/ml). Subsequent fractionation ofPSC into monomer-rich and oligomer-rich formulations was performed byadding an equal volume of 2× solubilization buffer (0.06 M phosphate,0.2 M NaCl, and 1.2 M glycerol, pH 7.0) with mixing. The solution wasleft to stand at 30° C. for 7 days.

At desired times, the solution was then centrifuged at 10,000 rpm and 4°C. for 10 minutes and decanted and the supernatant was retained. At eachtime point, the supernatant represented the monomer-rich collagenfraction and the pellet represented the oligomer-rich collagen fraction.The pellet was then resolubilized in 0.1 M acetic acid and theresolubilized pellet and the supernatant were dialyzed against 0.01 Macetic acid and lyophilized to dryness.

The composition of the oligomer-rich and monomer-rich collagen fractionswere confirmed by sulfate polyacrylamide gel electrophoresis (SDS-PAGE)(4%) analysis (FIG. 19, Lanes 1-3) and Western blot analysis with acollagen α1(I) antibody (FIG. 19, Lanes 4-6). In FIG. 19, theunfractionated, PSC starting material is represented in Lanes 1 and 4,the oligomer-rich collagen fraction is represented in Lanes 2 and 5, andthe monomer-rich collagen fraction is represented in Lanes 3 and 6.

SDS-PAGE analysis showed that, in addition to the α1(I), α2(I), β11(I),β12(I), and γ(I) bands routinely observed in denatured purified collagenType I preparations, the PSC included a prominent band corresponding toa molecular weight of 260 kDa (Oligo 260) as well as high molecularweight (HMW) components with molecular weights greater than 300 kDa(FIG. 19, Lane 1). The β, Oligo260, γ, and HMW bands represent two ormore a chains that are covalently linked by natural collagen crosslinkchemistries, whereas the α1(I) and α2(I) bands, which are present at aratio of 2 to 1 respectively, represent component polypeptide chains(˜100 kDa) within a single triple helical collagen molecule. TheOligo260 and HMW components were retained at significant levels in theoligomer-rich fraction and found at substantially reduced levels in themonomer-rich fraction, suggesting that the Oligo260 and HMW componentsrepresent oligomer derivatives with intermolecular cross-links.

Western blot analysis used mouse monoclonal antibodies specific for TypeI (AB6308, Abcam, Cambridge, Mass.) and Type III (MAB 1343, Chemicon,Temecula) collagen. According to manufacturer's specification, theantibody for Type I collagen was developed against the full lengthnative protein. However, in the present study, this antibody showedspecificity to the α1(I) chain when applied to denatured collagens.Western blot analysis confirmed that α1(I), β11(I), β12(I), Oligo260,and HMW components contained the epitope for collagen α1(I) (FIG. 19,Lane 4).

Also, intrinsic viscosity measurements, which provide a measure of theAMW of collagen polymer solutions in their native, undenatured state,were performed. In brief, apparent viscosities of collagen solutions in0.01 N HCl were measured on an AR2000 rheometer (TA Instruments, NewCastle, Del.) with a cone geometry (40 mm, 2° cone angle). Viscositiesfor solutions representing collagen concentrations of 0.1-0.3 mg/ml weremeasured at shear rates of 100-1500 s⁻¹ at 10° C. The AMW was calculatedusing the Mark-Houwink equation, |η|=kM^(a), where a=1.8 and k wasdetermined for each shear rate assuming a monomer-rich AMW of 282 kDa.AMW was extrapolated to zero shear rate. Intrinsic viscositymeasurements indicated that monomer-rich fractions had an AMW value ofabout 282 kDa and oligomer-rich fractions had an AMW value of about603+/−92 kDa. These results confirmed the prominence of cross-linkedcollagen monomers within the oligomer-rich fraction. PSC yielded roughlyfour times more oligomer-rich collagen compared to monomer-richcollagen, based upon dry weight.

The monomer-rich and oligomer-rich fractions described above were alsorecombined at different ratios to create collagen formulations thatvaried in monomer/oligomer content and thus AMW.

Additional information regarding the separation of collagen intomonomer-rich and oligomer-rich fractions is set forth in U.S. PatentApplication Publication No. 2008/0268052 to Voytik-Harbin et al., theentire disclosure of which is expressly incorporated herein byreference, and is described in [1].

Polymerization of 3D Collagen Matrices

Matrices were polymerized as previously described [1]. Briefly ECFCswere seeded at 5×105 or 2×106 cells/ml into unpolymerized collagensolution and allowed to polymerize for 30 min at 37° C. and 5% CO2 andthen 120 μl of warm EBM-2 (Lonza, Basel, Switzerland) supplemented with40 ng/ml FGF and SRII [126] was added. Media was changed at 24 hours.For vacuole formation assays, matrices were set in duplicate and atleast three independent experiments were conducted.

Referring to FIGS. 5 A-C. Fibril density did not demonstrate a positivecorrelation with AMW, while G′ (FIG. 5B) and Ec (FIG. 5C) increasedlinearly with increasing AMW. Data represent mean±SD [1].

Toluidine Blue Staining of Collagen Matrices

For analysis of in vitro vacuole formation, cultures were fixed with 4%paraformaldehyde for 20 minutes at 48 hours. Scaffolds were stained with0.1% toluidine blue O in 30% methanol for 20 minutes and then washedwith PBS.

Analysis of In Vitro Vacuole Formation

A Leica DM IRE2 microscope (Leica Microsystems, Bannockburn, Ill.) withattached Retiga 4000R digital camera (QImaging, Surrey, BC, Canada) wasused to image the entire area of each matrix using a 10× objective andthe middle 9 fields of view were used for analysis. Lumens werequantified using a standard image analysis system Metamorph (MolecularDevices, Sunnyvale, Calif.). From these data average lumen area andtotal lumen area were calculated. Lumens were defined as areascompletely surround by a toluidine blue labeled endothelial cellmembrane.

Perturbation of ECFC Vacuole Formation

Chemical inhibitors TIMP-3 (5 μg/ml, R&D Systems, Minneapolis, Minn.),Casin (2.5 μM), NSC23766 (25 μM), or MAB17781 (1 μg/ml, R&D Systems,Minneapolis, Minn.) were added into the media during ECFC culture in 3.0mg/ml matrices polymerized with the oligomer rich fraction. Treatmentswere conducted in duplicate and at least three independent experimentswere conducted.

mRNA Isolation

After 48 hours of cultures, RNA was isolated from ECFCs seeded intocollagen matrices using a RNeasy Mini kit (Qiagen, Valenci, Calif.). Tenmatrices were pooled from each treatment group for an experiment. At thesame passage, RNA from ECFC grown on tissue culture plastic wasisolated. Three independent experiments were conducted. All RNA sampleswere quantified using a Nanodrop 1000 (Thermo Scientific). RNA qualitywas determined by the A260/A280 and A260/A230 ratios. Using reversetranscription cDNA was synthesized using SuperScript II ReverseTranscriptase (Invitrogen, Carlsbad, Calif.).

Quantitative Real Time RT-PCR (qRTPCR)

Quantitative real time RT-PCR was performed using custom designedprimers (Invitrogen, Carlsbad, Calif.) for the three genes of interest(Table 2), FastStart Universal SYBR Green Master (Roche) and a ABI7500Real-Time PCR system (ABI, Carlsbad, Calif.). Cycling conditions were asfollows: 95° C. for 10 minutes, followed by 40 cycles of 95° C. for 15seconds and 60° C. for 1 minute. 7500 Software (ABI, Carlsbad, Calif.)was used to determine the CT values. Data was analyzed using the 2-ΔCTmethod using ATP5B as a reference gene. Each sample was run intriplicate two separate times and a maximum standard deviation betweenCT values of triplicates of 0.3 was considered acceptable.

TABLE 2  Primer sequences for qRTPCR. Name Forward Primer Reverse PrimerATP5B CCACTACCAAGAAGGGAT GGGCAGGGTCAGTCAAGTC CTATCA (SEQ. ID NO. 2)(SEQ. ID NO. 1) Cdc42 CATCGGAATATGTACCGA TGCAGTATCAAAAAGTCCA CTGTTAGAGTA (SEQ. ID NO. 3) (SEQ. ID NO. 4) MT1- CTGTCAGGAATGAGGATCAGGGGTCACTGGAATGCTC MMP TGAA (SEQ. ID NO. 6) (SEQ. ID NO. 5) Rac1CTGATCAGTTACACAACC CATTGGCAGAATAATTGTC AATGC AAAGA (SEQ. ID NO. 7)(SEQ. ID NO. 8)

Western Blot

Protein was extracted from ECFCs seeded into collagen scaffolds after 48hours of culture. Total cellular extracts were electrophoresed by sodiumdodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE),transferred to nitrocellulose and probed with anti-FAK (H-1, sc-1688Santa Cruz Biotechnology, Santa Cruz, Calif.).

ECFC Transfection

Three FRET based Rho GTPases, RhoA, Rac 1, and Cdc42 plasmids, were usedfor these experiments. ECFCs were transfected by electroporation using aNeon Transfection Device (Invitrogen, Carlsbad, Calif.) following themanufacture's recommended parameters. Briefly, 1×107 cells/ml weretransfected using 1 μg/ml plasmid, a pulse voltage of 1000 mV, a pulsewidth of 20 ms, and a pulse number of 2.

Transfected ECFC Microscopic Imaging

Transfected ECFCs were imaged by DIC and fluorescence microscopy with aNikon microscope (Nikon, Tokyo, Japan) using a 40× oil immersionobjective. Excitation wavelengths for the cyan fluorescence probe (CFP)and the yellow fluorescence probe (YFP) were 475 nm and 535 nmrespectively. Images were processed using NIS elements imaging software(Nikon, Tokyo, Japan) using iterative deconvolution and automaticbackground subtraction to improve resolution and remove image noise.

Statistical Analysis

All values are presented as mean±standard error (SE). Analysis ofvariance was conducted using SAS statistical software (SAS Institute,Inc., Cary, N. C.) to determine if there was a significant effect of thevariable of interest on cell behavior. If a statistically significantrelationship was found Tukey's test for multiple comparisons was used todetermine which groups were significantly different or a two-tailedstudent's t-test was used to evaluate the significance between twogroups. Additionally, a Dunnett's post test was used to comparetreatment groups to a control group. A p-value of less than 0.05 wasconsidered significant.

Referring now to FIG. 6, Panels A-F, ECFCs were able to form vacuoles inall matrices tested when seeded into the matrices at a density of about,2×106 cells/ml. However, the number and size of the vacuoles varied withthe collagen concentration and cross link composition of the matrix.Previous reports from the literature have demonstrated an alteration incapillary morphogenesis with increasing matrix concentration in vitroand in vivo [74, 80, 122]. Consistent with these results collagenconcentration altered ECFC vacuole formation.

Referring now to FIGS. 9 A-C. ECFC Vacuole density and total vacuolearea demonstrated a significant increase from 8.43±1.34 to 15.00±4.62vacuoles/mm² and from 1519.34±312.66 to 7440.75±3431.11 μm² over therange of collagen concentrations tested, while vacuole area showed atrend of increasing with increasing collagen concentration from40.07±5.88 to 73.94±21.48 μm2 (FIG. 7 A-D). The increase in vacuole areawas due to a shift to right of the vacuole area distribution (FIG. 7C)and at the highest collagen concentration, vacuoles with an area greaterthan 500 μm² were found. The effect of cross link chemistry was alsoshown to have an effect on ECFC in vitro lumen formation.

Referring now to FIGS. 8 A-D. Matrices polymerized with the oligomerrich fraction demonstrated an increased vacuole density, vacuole area,and total vacuole area when compared to matrices polymerized with themonomer rich fraction. Vacuole density increased from 5.41±0.18 to16.81±2.38, vacuoles/mm²; vacuole area increased from 40.02±5.29 to71.15±13.03, μm²; and total vacuole area increased from 938.81±102.20 to6612.60±2187.77, μm² in matrices polymerized from monomer and oligomerrespectively. Additionally, there was a shift in the distribution ofECFC vacuole areas to the right in matrices polymerized from oligomerrich fractions compared to matrices polymerized from monomer richfractions. Thus, both collagen concentration and collagen cross linkchemistry alter ECFC vacuole formation in vitro.

Alteration in collagen cross link chemistry resulted in alteredvacuolization in matrices with matched shear storage modulus (stiffness)but different collagen concentrations. Matrices were polymerized fromboth the oligomer and monomer rich fractions with a shear storagemodulus of 200 Pa. Referring now to FIGS. 9 A-C, the matricespolymerized from the oligomer matrix demonstrated an increased vacuoledensity from 6.88±1.39 to 13.48±2.33 vacuoles/mm², and a trend ofincreased total vacuole area from 1455.69±210.37 to 3178.10±654.04 μm².The average vacuole area did not demonstrate a difference between the200 Pa stiffness matrices polymerized from oligomer rich and monomerrich fractions. These data suggest that at a given stiffness theoligomer rich matrices are better able to support ECFC derived vacuoles.It is important to note that while these matrices have the samestiffness they have different collagen concentrations. The monomer richmatrix has a collagen concentration of 2.79 mg/ml and the oligomer richmatrix has a collagen concentration of 1.55 mg/ml. These data suggestthat at a given stiffness a matrix with a reduced collagen concentrationis better able to support ECFC vacuole formation.

Referring now to FIGS. 9 D-F. Vacuole density and total vacuoledisplayed an increase with increasing AMW (increasing ratio of collagenoligomers to collagen monomers) at a fixed collagen concentration of 1.5mg/ml. To explore the role of the collagen cross link content inmatrices with a fixed collagen concentration oligomer and monomer richfractions were mixed to yield matrices with varied oligomer and monomerratios. Matrices were polymerized with a collagen concentration of 1.5mg/ml and a collagen oligomer to monomer ratio of 0:100, 50:50, and100:0. Vacuole density demonstrated an increase from 5.04±0.20 to14.75±1.58 vacuoles/mm² and total vacuole area increased from1180.96±18.99 to 2090.31±51.72 μm² with increasing AMW, while averagevacuole area did not demonstrate a significant difference with AMWranging from 53.56±2.29 to 51.91±5.31 μm². These data further supportthe role for the collagen cross link chemistry as an important modulatorof capillary morphogenesis.

Referring now to FIG. 6, ECFCs seeded into collagen based scaffoldsundergo capillary morphogenesis. ECFCs seeded into matrices polymerizedfrom oligomer rich fractions at 0.5 (FIG. 6 panel A), 1.5 (FIG. 6 panelB), and 3.0 (FIG. 6 panel C) mg/ml and monomer rich fractions at 0.5(FIG. 6 panel D), 1.5 (FIG. 6 panel E), and 3.0 (FIG. 6 panel F) mg/mlformed vacuoles of different sizes and at different densities (scalebare=100 μm).

Referring now to FIGS. 7 A-C, collagen levels alter ECFC in vitrovacuole formation in collagen based scaffolds. ECFC vacuole density(FIG. 7A) and total vacuole area (FIG. 7D) significantly increased withincreasing collagen concentration. Vacuole area (FIG. 7B) showed a trendof increasing with increasing collagen concentration due to a shifttoward larger vacuole areas (FIG. 7C) (asterisk denotes p<0.05).

Referring now to FIGS. 10 A-C, matrices polymerized from oligomer richfractions show an increase in vacuole density and total vacuole areaover a range of collagen concentrations from 0.5 mg/ml to 3.0 mg/ml.Matrices were polymerized from either oligomer or monomer rich fractionsat 0.5, 1.5, and 3.0 mg/ml. At each collagen concentration the matrixpolymerized from the oligomer rich fraction demonstrated an increasedvacuole density and total vacuole area. Additionally, at 3.0 mg/ml thematrix polymerized from the oligomer rich fraction displayed anincreased average vacuole area compared to the matrix polymerized fromthe monomer rich fraction. The vacuole area distribution displayed ashift toward vacuoles of larger size for oligomer rich matrices with acollagen concentration of 3.0 mg/ml compared to 3.0 mg/ml matricespolymerized from monomer rich fraction (FIG. 10F). This alteration invacuole area distribution with altered oligomer content is not seen atlower collagen concentrations.

Referring now to FIGS. 8 A-D, collagen matrices polymerized witholigomer and monomer rich fractions demonstrate altered ECFC in vitrovacuole formation. ECFCs displayed increased vacuole density (FIG. 8A),vacuole area (FIG. 8B), and total vacuole area (FIG. 8D) in matricespolymerized from oligomer rich fractions compared to those polymerizedfrom monomeric rich fractions. The percentage of vacuoles with an areagreater than 100 μm² was increased in the oligomer rich scaffolds (FIG.8C, asterisk denotes p<0.05). Data represent Mean±SE and n=3.

Referring now to FIGS. 9 A-F, alterations in collagen cross link and AMWalter ECFC vacuole formation. Matrices polymerized from oligomer andmonomer rich fraction with a stiffness of 200 Pa display differentialECFC vacuole formation (A-C). Matrices polymerized at 1.5 mg/ml withvaried AMW demonstrate an increase in vacuole density and total vacuolearea with increasing AMW (D-F, asterisk denotes p<0.05). Data representMean±SE and n=3.

Characterization of Matrix-Integrin-Cytoskeleton Signaling Axis in ECFCVacuolization

Capillary morphogenesis has been shown to be dependent upon signalingthrough 131 integrin to Rho GTPase family members Cdc42 and Rac1 [127]and to also depend on matrix remodeling by MT1-MMP [128]. To firstdetermine if this signaling axis was involved in ECFC vacuole formationchemical inhibitors to β1 integrin, Cdc42, Rac1, and MT1-MMP were addedto cultures to test if they would alter cell behavior.

Referring now to FIGS. 11 A-H, inhibition of the β1 integrin, Cdc42,Rac1, and MT1-MMP activity decreases ECFC total vascular area. Chemicalinhibitors MAB17781, Casin, NSC23766 [129], and TIMP-3 were individuallyadded to the cultures to inhibit β1 integrin, Cdc42, Rac1, and MT1-MMPactivity respectively. While all inhibited ECFC vacuole formation,MAB17781 and TIMP-3 had the greatest reduction in vacuole density from11.24±0.65 to 2.71±0.46 and 3.31±1.38 vacuoles/mm² respectively, vacuolearea from 536.09±138.24 to 259.60±67.08 and 146.04±45.86 μm²respectively, and total vacuole area from 19103.60±5447.67 to2387.82±620.56 an 2288.76±1685.92 μm² respectively. Additionally,inhibition of Cdc42 signaling decreased total vacuole area by decreasingvacuole density whereas inhibition of Rac1 signaling decreased totalvacuole area by decreasing average vacuole area (FIGS. 11 G, H). Theseresults suggest that while all of the proteins are involved in ECFCvacuole formation they play a nuanced role in the process.

Referring now to FIGS. 10 A-F, varied collagen cross link compositionresults in differential ECFC vacuole formation in vitro over a range ofcollagen concentrations. At 0.5, 1.5, and 3.0 mg/ml, matricespolymerized from oligomer rich fractions displayed an increase invacuole density (FIG. 10A) and total vacuole area (FIG. 10C). At 3.0mg/ml vacuole area was also significantly increased in oligomer richscaffolds (FIG. 10B). This was due to a shift in the vacuole areadistributions (FIGS. D-F, asterisk denotes p<0.05).

Alteration in mRNA expression by altered matrix microenvironment Cultureof ECFCs in a 3D collagen matrix alters mRNA expression of Cdc42, Rac1,and MT1-MMP compared to ECFC grown in 2D culture. To begin to test fordifferences in cell signaling, mRNA was isolated from ECFCs grown on 2Dtissue culture plastic and from ECFCs cultured in collagen matrices. ThemRNA expression levels for Cdc42, Rac1, and MT1-MMP were normalized toATP5B, a housekeeping gene for cells grown in 2D and 3D environments.Transcript levels were increased in ECFCs grown in a 3D environmentcompared to ECFCs cultured in a 2D environment for Cdc42 from0.263±0.128 to 0.475±0.03, Rac1 from 0.374±0.069 to 0.840±0.093, andMT1-MMP from 0.012±0.001 to 0.308±0.033 (FIG. 12C). These data are notsurprising given the differences in cell.

Referring now to FIGS. 11 A-H, alteration in signaling through thematrix integrin cytoskeleton axis disrupts ECFC in vitro vacuoleformation. (A) control cultures displayed normal ECFC vacuole formation.The addition of MAB17781 (B), Casin (C), NSC23766 (D), or TIMP-3 (E,scale bare=100 μm) disrupt vacuole formation and lead to reductions invacuole density (F), vacuole area (G), and total vacuole area (H,asterisk denotes p<0.05 compared to control cultures).

Referring now to FIG. 12 A-B, culture of ECFCs in 3D collagen basedscaffolds alters mRNA expression. Referring to FIG. 12 A, ECFCsdisplayed different morphology when grown on tissue culture plastic(FIG. 12 A, panel A, scale bar=100 μm) and in collagen matrices (FIG. 12A, panel B, scale bar=100 μm) and exhibited altered expression of Cdc42,Rac1, and MT1-MMP mRNA (C, asterisk denotes p<0.05). Culture of ECFCs in3D collagen based scaffolds alters mRNA expression. ECFCs displayeddifferent morphology when grown on tissue culture plastic (FIG. 12 A,panel A, scale bar=100 μm) and in collagen matrices (FIG. 12 A, panel B,scale bar=100 μm) and exhibited altered expression of Cdc42, Rac1, andMT1-MMP mRNA (FIG. 12 B, asterisk denotes p<0.05). Data representMean±SE and n=3. morphology in the two different culture environmentsand further stresses the importance of investigating cell behavior in a3D context which better recapitulates the in vivo environment. ECFCsseeded into matrices polymerized from oligomer rich fractions exhibitincreased mRNA expression of Cdc42 and MT1-MMP and decreased expressionof Rac1 compared to monomer rich fractions.

Next the mRNA expression from ECFCs cultured in matrices polymerizedfrom both oligomer and monomer rich fractions at 0.5 and 3.0 mg/ml werecompared. Referring now to FIGS. 13 A-C, collagen cross link chemistryalters ECFC mRNA expression. ECFC mRNA expression of Cdc42 (FIG. 13A),Rac1 (FIG. 13B), and MT1-MMP (FIG. 13C) in matrices polymerized withmonomer or oligomer rich fractions at 0.5 and 3.0 mg/ml is altered.Matrices polymerized with oligomer rich collagen resulted in an upregulation of Cdc42 and MT1-MMP expression and a down regulation of Rac1compared with monomer rich matrices (D, asterisk denotes p<0.05). Whilethere was a trend of increased transcript levels for Cdc42 and MT1-MMPand decreased Rac1 mRNA expression in oligomer rich matrices at 0.5 and3.0 mg/ml compared to monomer rich matrices at the same collagenconcentration, the only significant difference was seen between 0.5mg/ml oligomer rich and monomer rich matrices with expression levels of0.571±0.022 and 0.393±0.054 respectively.

However, in comparing oligomer rich and monomer rich matrices, therewere significant differences in mRNA expression in all three genes (FIG.13D). Still referring to FIG. 13 D, the expression levels in oligomerand monomer matrices of Cdc42 mRNA was 0.535±0.021 and 0.415±0.039, theexpression levels of Rac 1 mRNA in oligomer and monomer matrices was0.607±0.07 and 1.074±0.108, and the expression of MT1-MMP mRNA was0.378±0.048 and 0.239±0.025 respectively. These data suggest that crosslink chemistry plays a role in regulating ECFC mRNA expression duringcapillary morphogenesis.

Alteration in Protein Expression by Altered Scaffold BiophysicalProperties

To determine if the varied microenvironment that the ECFCs experiencedaltered the protein expression level of ECFCs seeded into matricespolymerized of monomer and oligomer rich collagen, protein was isolatedfrom ECFCs seeded into collagen matrices after 48 hours of culture.Endothelial cells interact with the collagen fibrils via integrinbinding and activation leading to integrin clustering and focal adhesionformation. This outside in signaling is modulated by alterations in thebiophysical properties of the matrix. To begin to interrogatealterations in the outside in signaling, focal adhesion kinase (FAK)protein expression was determined.

Referring now to FIG. 14, FAK expression is elevated in ECFCs seededinto matrices polymerized from oligomer rich fractions compared tomatrices polymerized from monomer rich fractions. ECFCs seeded intooligomer (O) or monomer (M) rich matrices at matched stiffness orcollagen concentration display increased total FAK expression.

These data suggest that FAK protein expression is elevated in ECFCsseeded into matrices polymerized from oligomer rich fractions comparedto ECFCs seeded into matrices polymerized from monomer rich fractions(FIG. 14). This elevation in FAK expression suggests increased outsidein signaling into the endothelial cells which could be partiallyresponsible for the increased vacuole formation in these oligomer richmatrices.

RhoGTPases are Present at Site of ECFC Vacuole Formation

To begin to determine the alteration in RhoGTPase activity duringcapillary morphogenesis, ECFC were transfected with FRET based RhoA,Rac1, or Cdc42 plasmids and then seeded into oligomer rich matrices.Referring now to FIG. 15, RhoGTPases RhoA, Rac 1, and Cdc42 are activeat the site of vacuole formation in ECFCs (Images are at 400×). Allthree proteins appear to be active at the site of vacuole formation inECFCs further suggesting they each may play a role in capillarymorphogenesis.

Cell Density Alters ECFC Vacuole Formation in Collagen Matrices

The ratio of apparent matrix stiffness to cell traction has beendemonstrated in the literature to impact capillary morphogenesis [74,81]. In these previous reports when the ratio was too high capillarymorphogenesis failed to occur, but when it was too low excessivecontraction of the matrix resulted in cell death [74]. Referring now toFIGS. 16 A-C, ECFC density alters vacuole formation in collagenmatrices. Increased cell seeding density results in increased vacuoledensity (FIG. 16 A), but decreased vacuole area (FIG. 16B), and nochange in total vacuole area (FIG. 16C, asterisk denotes p<0.05). Totest for the effect of alterations in cell traction, the cell density ofECFCs seeded into the collagen based scaffolds was reduced from 2×106cells/ml to 5×105 cells/ml. ECFCs seeded at a lower cell densitydemonstrated a lower vacuole density of 5.80±0.49 compared to 10.62±1.27vacuoles/mm² at the higher cell seeding density (FIG. 16A), but anincreased vacuole area of 94.90±8.43 compared to 54.08±5.27 μm² (FIG.16B). Further the total vascular area was not significantly differentbetween the two seeding densities.

Matrices polymerized from oligomer and monomer rich fractions responddifferently to alterations in cell seeding density. Matrices werepolymerized from oligomer or monomer rich fractions with a collagenconcentration of 0.5 and 3.0 mg/ml and seeded with either 5×10⁵ or 2×10⁶cells/ml. Referring now to FIGS. 17 A-C, cell density alters ECFC invitro vacuole formation. Increasing cell density resulted in increasedvacuole density (FIG. 17A) for all matrices, but decreased vacuole area(FIG. 17B) and total vascular area (FIG. 17C) for all matrices except3.0 mg/ml oligomer rich scaffolds (asterisk denotes p<0.05). As expectedan increase in cell density resulted in an increase in vacuole densityfor each matrix (FIG. 17A). However the average vacuole area decreasedwith increasing cell density for all matrices except the oligomer richmatrix polymerized at 3 mg/ml (FIG. 17B). As a result total vacuole areadecreased or remained unchanged with an increase in cell density for allmatrices except for the 3.0 mg/ml oligomer matrix (FIG. 17C). Thisunexpected finding is due to a shift in the distribution of ECFCvacuoles toward smaller vacuole areas in the monomer rich and 0.5 mg/mloligomer rich matrices (FIGS. 18 A-C). In contrast there is no leftwardshift of the vacuole distribution in the 3.0 mg/ml oligomer richscaffold and a trend of rightward shifting of vacuole size in thismatrix. These data suggested the possibility that the ECFC generatedtraction and matrix remodeling could not be sufficiently resisted by themonomer rich and 0.5 mg/ml oligomer rich scaffolds.

Alterations in ECFC cell density result in varied vacuole areadistributions. Referring now to FIGS. 18 A-D, the distribution ofvacuole areas was shifted leftward for monomer rich matrices polymerizedwith a collagen concentration of 0.5 (FIG. 18A) and 3.0 (FIG. 18B) mg/mland for oligomer rich matrices polymerized with a collagen concentrationof 0.5 mg/ml (FIG. 18C). However this shift is not seen in oligomer richmatrices polymerized at a collagen concentration of 3.0 mg/ml (FIG.18D).

For the purposes of promoting an understanding of the principles of thenovel technology, reference will now be made to the preferredembodiments thereof, and specific language will be used to describe thesame. It will nevertheless be understood that no limitation of the scopeof the novel technology is thereby intended, such alterations,modifications, and further applications of the principles of the noveltechnology being contemplated as would normally occur to one skilled inthe art to which the novel technology relates are within the scope ofthis disclosure and the claims.

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We claim:
 1. A vessel forming matrix comprising: a population ofendothelial colony forming cells in a polymerized matrix comprising atleast one fraction of an acid soluble Type I collagen.
 2. The matrixaccording to claim 1, wherein said at least one fraction of the acidsoluble Type I collagen is selected from the group consisting ofmonomers having an average molecular weight of about 282 kDa andoligomers having an average molecular weight of about 603 kDa.
 3. Thematrix according to claim 1, wherein said at least one fraction of acidsoluble Type I collagen includes a telopeptide reactive aldehyde.
 4. Thematrix according to claim 1, further comprising a second fraction of theacid soluble Type I collagen that differs from the at least onefraction.
 5. The matrix according to claim 4, wherein the at least onefraction is an oligomer-rich fraction and the second fraction is amonomer-rich fraction.
 6. The matrix according to claim 5, wherein thematrix comprises more of the oligomer-rich fraction than themonomer-rich fraction.
 7. The matrix according to claim 1, wherein saidat least one fraction of acid soluble Type I collagen is present in thematrix at a concentration of about 0.5 to about 3.0 mg/ml.
 8. The matrixaccording to claim 1, wherein said population of endothelial colonyforming cells is present in the matrix at a concentration of about 5×10⁵to about 2×10⁶ cells/ml.
 9. A method of promoting vessel formationcomprising the steps of: obtaining an engineered collagen matrix from anoligomer-rich fraction of a Type I acid soluble collagen; and seedingthe matrix with a plurality of endothelial colony forming cells to formone or more vessels within the matrix.
 10. The method according to claim9, wherein the matrix is richer in Type I acid soluble collagenoligomers than in Type I acid soluble collagen monomers.
 11. The methodaccording to claim 9, wherein the average molecular weight of thecollagen in the matrix is about 603 kDa.
 12. The method according toclaim 9, wherein the average molecular weight of the collagen in thematrix is greater than about 282 kDa.
 13. The method according to claim9, wherein the matrix has a collagen concentration of about 0.5 to about3.0 mg/ml.
 14. The method according to claim 9, wherein the obtainingstep comprises isolating oligomers from the Type I acid solublecollagen.
 15. The method according to claim 9, wherein the matrix isstiffer than the Type I collagen.
 16. The method according to claim 9,wherein the matrix is in an unpolymerized state during the seeding step.17. The method according to claim 9, wherein the seeding step comprisesseeding the matrix with about 5×10⁵ to about 2×10⁶ cells/ml.
 18. Themethod according to claim 9, further comprising the step of implantingthe matrix into a diseased area of a patient.
 19. The method accordingto claim 18, wherein the diseased area of the patient comprises anischemic limb.
 20. The method according to claim 9, wherein theobtaining step comprises independently controlling a stiffness of thematrix and a collagen concentration of the matrix.